measuring dna mechanics on the genome scale · 2020. 8. 17. · dna sequences that are more bent8...

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Measuring DNA mechanics on the genome scale

Aakash Basu1, 2, Dmitriy G. Bobrovnikov1, Zan Qureshi3, Tunc Kayikcioglu1, 4, Thuy T. M. Ngo4, Anand

Ranjan5, Sebastian Eustermann6, 7, Basilio Cieza3, Michael T. Morgan1, Miroslav Hejna2, 8, H. Tomas Rube2,

8, Karl-Peter Hopfner6, 7, Cynthia Wolberger1, Jun S. Song2, 8, 9 and Taekjip Ha1, 2, 3, 10, 11, a

1Department of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine,

Baltimore, MD 21205, USA; 2Department of Physics, University of Illinois at Urbana Champaign, Urbana,

IL 61801, USA; 3Department of Biophysics, Johns Hopkins University, Baltimore, MD 21218, USA;

4Center for Biophysics and Computational Biology, University of Illinois at Urbana-Champaign, Urbana,

IL 61801, Illinois, USA; 5Department of Biology, Johns Hopkins University, Baltimore, MD 21218, USA;

6Department of Biochemistry, Ludwig-Maximilians-Universität, Feodor-Lynen-Strasse 25, 81377 Munich,

Germany; 7Gene Center, Ludwig-Maximilians-Universität, Feodor-Lynen-Strasse 25, 81377 Munich,

Germany; 8Carl R. Woese Institute for Genomic Biology, University of Illinois at Urbana-Champaign,

Urbana, IL, USA; 9Cancer Center at Illinois, University of Illinois, Urbana, IL 61801, USA; 10Department

of Biomedical Engineering, Johns Hopkins University, Baltimore, MD 21205; 11Howard Hughes Medical

Institute, Baltimore, MD 21205, USA.

aTo whom correspondence should be addressed (e-mail: [email protected])

Abstract

Mechanical deformations of DNA such as bending are ubiquitous and implicated in diverse cellular

functions1. However, the lack of high-throughput tools to directly measure the mechanical properties of

DNA limits our understanding of whether and how DNA sequences modulate DNA mechanics and

associated chromatin transactions genome-wide. We developed an assay called loop-seq to measure the

intrinsic cyclizability of DNA – a proxy for DNA bendability – in high throughput. We measured the

(which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. The copyright holder for this preprintthis version posted August 18, 2020. ; https://doi.org/10.1101/2020.08.17.255042doi: bioRxiv preprint

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intrinsic cyclizabilities of 270,806 50 bp DNA fragments that span the entire length of S. cerevisiae

chromosome V and other genomic regions, and also include random sequences. We discovered sequence-

encoded regions of unusually low bendability upstream of Transcription Start Sites (TSSs). These regions

disfavor the sharp DNA bending required for nucleosome formation and are co-centric with known

Nucleosome Depleted Regions (NDRs). We show biochemically that low bendability of linker DNA

located about 40 bp away from a nucleosome edge inhibits nucleosome sliding into the linker by the

chromatin remodeler INO80. The observation explains how INO80 can create promoter-proximal

nucleosomal arrays in the absence of any other factors2 by reading the DNA mechanical landscape. We

show that chromosome wide, nucleosomes are characterized by high DNA bendability near dyads and low

bendability near the linkers. This contrast increases for nucleosomes deeper into gene bodies, suggesting

that DNA mechanics plays a previously unappreciated role in organizing nucleosomes far from the TSS,

where nucleosome remodelers predominate. Importantly, random substitution of synonymous codons does

not preserve this contrast, suggesting that the evolution of codon choice has been impacted by selective

pressure to preserve sequence-encoded mechanical modulations along genes. We also provide evidence

that transcription through the TSS-proximal nucleosomes is impacted by local DNA mechanics. Overall,

this first genome-scale map of DNA mechanics hints at a ‘mechanical code’ with broad functional

implications.

Measuring DNA mechanics in high throughput

DNA looping (or cyclization) assays have long been used to measure DNA bendability3,4. Recently,

a single molecule Fluorescence Resonance Energy Transfer5 (smFRET)-based DNA looping assay was

developed6, whereby looping of a ~100 basepair (bp) DNA duplex flanked by complementary 10 nucleotide

(nt) single-stranded overhangs is detected via an increase in FRET between fluorophores located at each

end of DNA (Fig. 1a). The looping rate thus obtained has been interpreted as a measure of DNA bendability.

In this assay, chemically synthesized single strands of DNA had to be annealed directly without PCR

amplification to generate a duplex region flanked by long 10 nt overhangs. We simplified the process by


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developing a nicking-based method that allows the in situ conversion of a 120 bp duplex DNA (which can

be produced via PCR amplification) into a 100 bp duplex flanked by 10 nt single-stranded overhangs (Fig.

1b). Using FRET6, we measured the looping times of ten DNA fragments with different sequences

(Supplementary Note 1). The looping times varied by more than an order of magnitude (Fig. 1c), confirming

the previously reported result that DNA sequence can have a profound effect on DNA looping at the ~100

bp length scale6,7. However, looping assays and all previous methods to directly measure DNA bendability

have limited throughput, which greatly limits our knowledge of how DNA mechanics is modulated by

sequence, varies along genomes, and influences chromosome transactions.

Systematic Enrichment of Ligands by Exponential Enrichment (SELEX) has been used to enrich

DNA sequences that are more bent8 or more loopable9 through many rounds of selection of rapidly looping

DNA from a vast random pool of loopable molecules and PCR amplification of selected molecules. These

assays revealed, for example, certain periodic dinucleotide distributions found in the variable regions of

highly loopable DNA. However, direct bendability measurements of specified sequences of interest, such

as those than span genomic regions, have never been reported in high throughput.

In order to extend direct DNA bendability measurements to a much larger sequence space, we

established a sequencing-based approach termed loop-seq, which builds on previous low-throughput single-

molecule looping6 and SELEX selection methods9. Using the nicking approach, we generated a library of

up to ~90,000 different specified template sequences immobilized on streptavidin-coated beads. Library

members had a central 50 bp duplex region of variable sequence flanked by 25 bp left and right duplex

adapters and 10 nt single-stranded complementary overhangs (Fig. 1d). Looping was initiated in high salt

for 1 minute, after which unlooped DNA molecules were digested with an exonuclease9 (RecBCD) that

requires free DNA ends, thus preserving the looped molecules. The enriched library was sequenced, and

the cyclizability of each sequence was defined as the natural logarithm of the ratio of the relative population

of that sequence in the enriched library to that in an identically treated control in which only the digestion

step was omitted (Fig. 1e, Supplementary Note 2).


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The looping times of the 10 sequences determined via smFRET (Fig. 1c) were strongly anti-

correlated with their cyclizability values obtained by performing loop-seq on a large library containing

those 10 (along with 19,897 other) sequences (Fig. 1f). This confirmed that cyclizability is a good measure

of looping rate. Additionally, varying the time for which looping is permitted before RecBCD digestion

allowed for the measurement of the full looping kinetic curves of all sequences in the library (‘Timecourse

loop-seq’, Supplementary Note 3 and Extended Data Fig. 2). The looped population could comprise closed

structures with alternate shapes and basepairing geometries10,11 (Extended Data Fig. 1). However,

irrespective of looped geometry, control experiments indicate that most looped molecules are protected

from RecBCD digestion, and also serve to validate several other aspects of the assay (Extended Data Fig.

3).

We found that the distance of the biotin tether from the end of each molecule (n, Fig. 1d) imposed

an oscillatory modulation on cyclizability, possibly owing to a sequence-dependent preference for the

rotational orientation of the biotin tether (Supplementary Note 7). By varying n and performing loop-seq

multiple times, we measured the mean, amplitude, and phase associated with this oscillation for every

library sequence. We called the mean term the “intrinsic cyclizability” and showed that it is independent of

the tethering geometry and rotational phasing (Supplementary Notes 7-8, Extended Data Fig. 4). Both

dynamic flexibility and static bending may contribute to intrinsic cyclizability. Regardless of interpretation,

intrinsic cyclizability is a measurable mechanical property that can be compared to functional properties of

chromosomal DNA.

DNA at yeast NDRs is rigid

We used loop-seq to query the role of DNA mechanical properties in establishing characteristic

features of genes that regulate expression, such as Nucleosome Depleted Regions (NDRs) upstream of the

Transcription Start Sites (TSSs) and well-ordered arrays of downstream nucleosomes positioned at

characteristic distances from the TSSs12. Although several lines of evidence had suggested that DNA

mechanics, in addition to transcription factors and chromatin remodelers2,13,14, plays a role in this regard by


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modulating nucleosome organization2,9,15,16, the mechanical properties of DNA along promoters and genes

have never been directly measured. We measured the intrinsic cyclizabilities of DNA fragments (‘Tiling

Library’, Supplementary Note 9) that tile the region from 600 bp upstream to 400 bp downstream of the +1

nucleosome dyads of 576 genes in S. cerevisiae at 7 bp resolution (Fig. 2a). We discovered a sharply defined

region of rigid DNA (i.e. with unusually low intrinsic cyclizability) located in the NDR17 (Fig. 2b). Further,

there are many genes where our measurements are sensitive enough to detect this region of high rigidity

without the need to average across multiple genes (Fig. 2c, Extended Data Fig. 5). As nucleosome assembly

requires extensive DNA bending, the low intrinsic cyclizability of DNA around the NDR is likely to favor

nucleosome depletion.

Chromatin remodelers sense DNA mechanics

Chromatin remodelers have been proposed to be critical in establishing the well-ordered array of

nucleosomes downstream of TSSs by stacking nucleosomes against a barrier just upstream of the TSS14.

What could constitute such a barrier has been a matter of debate, and transcription factors13 and paused

polymerases18 have been suggested to contribute. Notably, in vitro chromatin reconstitution experiments2

showed that the remodeler INO80 can both position the +1 (and -1) nucleosomes and establish the NDRs

in S. cerevisiae even in the absence of any such factors. We therefore asked whether the sequence-encoded

rigid DNA region in the NDR can contribute to nucleosome positioning near promoters by serving as a

barrier to the sliding activities of INO80.

To effect sliding, INO80 requires at least 40 – 50 bp of free extranucleosomal DNA ahead of the

nucleosome19,20. The region around 40 – 50 bp ahead of the sliding nucleosome’s edge is enganged by the

Arp8 module of INO8021–23, and disrupting the module’s DNA binding via mutation abolishes sliding and

reduces +1 positioning genome-wide23. Intriguingly, we found that the region of rigid DNA also starts ~43

bp upstream of the edge of the +1 nucleosome (Fig. 2b). This would place the Arp8 module in contact with

highly rigid DNA if INO80 were to slide the +1 nucleosome upstream from its canonical position (Fig. 2d).


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If highly rigid DNA interferes with Arp8 module binding, further upstream sliding of the +1 nucleosome

would be hindered, helping position the +1 nucleosome and define the NDR.

To directly test the role of upstream DNA rigidity in +1 nucleosome positioning by INO80, we

biochemically measured the effect of rigid DNA located ~40 bp ahead of a nucleosome on sliding by

INO80. Using gel shift, we assayed sliding of nucleosomes formed on the 147 bp 601 sequence into adjacent

80 bp linkers. We chose three pairs of such constructs, each containing one construct with a linker that was

uniformly flexible and another with a linker that had a significantly more rigid region near the middle

(Supplementary Note 11, Extended Data Fig. 6a). In all three pairs, the extent of sliding (Supplementary

Note 11) was lower for the nucleosome formed on the construct with the rigid linker (Fig. 2f, Extended

Data Figs. 6b-c, 7). Various factors could cause this reduced sliding (Supplementary Note 11). Regardless,

the observation is consistent with a model where the rigid DNA region starting ~43 bp upstream of the

canonical +1 nucleosome’s edge (Fig. 2b) serves as a barrier that hinders further upstream sliding of the +1

nucleosome by INO80, possibly aided by other barriers set up by factors such as RSC, gene regulatory

factors, and transcription factors2,13. Structural details behind rigidity sensing by the Arp8 module must

await future studies.

DNA mechanics impacts nucleosome organization

As nucleosomes involve extensive DNA bending, we asked if modulations in intrinsic cyclizability

may directly contribute to nucleosome organization, in addition to the stacking action of remodelers14.

Indeed, DNA at the canonical dyad locations of the +/-1 nucleosomes, and to a lesser extent the +2, +3 and

+4 nucleosomes, have significantly higher intrinsic cyclizability than surrounding DNA (Fig. 2b).

Consistent with this observation, promoters classified as having a fragile -1 nucleosome24 have more rigid

DNA at the location of the -1 nucleosome (Fig. 3a).

Several earlier studies have shed light on the role of DNA mechanics in nucleosome formation25.

The fact that bendable DNA forms good substrates for nucleosomes and vice versa, has been demonstrated

for various selected sequences4,26–30. Further, DNA selected for high loopability from a large random pool


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possess periodic distribution in dinucleotide contents9, which is also a feature found in ~3% of native yeast

nucleosomal sequences31. However, the mechanical properties of known nucleosomal DNA sequences have

never been directly measured in high throughput. To achieve this for nucleosomes along an entire yeast

chromosome, we measured intrinsic cyclizability along S. cerevisiae chromosome V at 7 bp resolution

(‘ChrV Library’, Supplementary Note 12, Extended Data Fig. 8). We first confirmed that intrinsic

cyclizability shows the characteristic pronounced dip around the NDR when averaged over the 227 genes

along chromosome V that have both ends mapped with high confidence25 (Fig. 3b). We found that

chromosome-wide, DNA at nucleosomal dyad locations tends to have significantly higher intrinsic

cyclizability than the surrounding linker DNA (Fig. 3c), suggesting that sequence-dependent modulations

in DNA mechanics contribute to global nucleosome organization. We also found that nucleosomes are

better positioned in vivo on more intrinsically cyclizable DNA (Fig. 3c, Extended Data Fig. 9a-c). Among

TSS proximal nucleosomes, the correlation is strongest for +1 nucleosomes (Fig. 3d, Extended Data Fig.

9d-e).

Among nucleosomes that lie along transcribed regions, TSS-distal nucleosomes have a higher

intrinsic cyclizability contrast between the dyad and the edges than TSS-proximal nucleosomes (Figs. 3e-

f). This was contrary to expectation because TSS-proximal nucleosomes are known to be better positioned

than TSS-distal nucleosomes14,32. TSS-proximal nucleosomes are likely primarily organized by chromatin

remodelers into ordered arrays via stacking against the NDR barrier2,14. However, beyond the +4

nucleosome, the stacking effect has been shown to dissipate14, whereas our data shows that modulations in

intrinsic cyclizability become more prominent (Fig. 3e-f). Thus nucleosomes that lie deeper in gene-bodies

may rely more on sequence-encoded intrinsic cyclizability modulations for positioning.

Codon choice optimizes mechanical modulations

We next asked whether the strong modulation in intrinsic cyclizability for nucleosomes deep in the

gene body would be preserved if the sequences were altered by using alternate codons that code for the

same amino acids. We selected 500 +7 nucleosomes in S. cerevisiae and generated four sets of codon-


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altered sequences spanning the region around these nucleosomes, while preserving the amino acid

sequences encoded. The natural codon usage frequency was considered when choosing synonymous codons

in the first two sets, and was ignored in the next two (supplementary note 14). By performing loop-seq

(‘Library L’, supplementary note 14), we measured intrinsic cyclizability at 7 bp resolution in the 200 bp

region flanking the 500 +7 nucleosome dyads and their codon-altered sequences. Native sequences have a

characteristic intrinsic cyclizability pattern – high near the dyads and low near the edges – which is absent

in the four codon-altered sets (Fig. 3g). Thus naturally occurring codons are optimized to establish

sequence-dependent intrinsic cyclizability modulations along genes that are favorable to the organization

of gene body nucleosomes, suggesting that the evolution of codon choice in S. cerevisiae has been impacted

by a selective pressure to preserve such modulations. The observation also points to a hitherto unappreciated

significance of positioning nucleosomes that lie deeper in the gene body.

TSS-proximal nucleosomes are asymmetric

Several critical processes such as transcription and DNA replication require nucleosome

unraveling. DNA could potentially peel off from either end, in a manner modulated by bendability. Indeed

asymmetry in DNA bendability across the 601 nucleosome leads to asymmetric unraveling under tension33.

Biochemical analysis has shown that yeast RNA polymerase II transcribing a 601 nucleosome produces

four times more full-length transcripts when it enters the nucleosome through the ‘TA-rich’ side that

contains the phased TA repeats34. Using loop-seq, we also found that the TA-rich side has significantly

higher intrinsic cyclizability (Fig. 4a, Supplementary Note 18). This observation is consistent with an idea

that RNA polymerase might better negotiate with a nucleosomal barrier when it first interacts with the side

of the nucleosome containing DNA with higher intrinsic cyclizability. We constructed a library containing

the 50 bp DNA fragments immediately to the left and right of the dyads of ~10,000 well-positioned S.

cerevisiae nucleosomes (‘Cerevisiae Nucleosomal Library’, Supplementary Note 4, Fig. 4b). We found

that DNA at well-occupied +1 and +2 nucleosomes indeed has, on average, higher intrinsic cyclizability on

the promoter-proximal face than the distal face (Fig. 4c), thereby suggesting that this asymmetry may favor


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polymerase translocation. Consistently, this asymmetry is accentuated among the highly expressed genes

and absent among poorly expressed genes (Fig. 4d).

Conclusions

Intrinsic cyclizability is, thus far, the only mechanical property of DNA to be directly measured in

high throughput, and will likely aid our understanding of how DNA mechanics influences chromatin

transactions involving diverse factors such as topoisomerases, transcription factors, polymerases, structure

maintenance of chromatin proteins and so on. The large dataset enabled by loop-seq should make it possible

to develop comprehensive models to predict intrinsic cyclizability and other physical properties from DNA

sequence. Preliminary analysis showed that simple sequence features such as GC content, polyA tracts, and

dinucleotide parameters are generally poor or insufficient predictors of intrinsic cyclizability

(Supplementary Note 16, Extended Data Fig. 10).

Our measurements suggest that intrinsic cyclizability is functionally important and must have

applied selective pressure throughout the evolution of genomes. It remains to be investigated how genetic

information content and the mechanical properties of DNA are linked, and how the sequence-dependent

mechanical response of DNA to molecular-scale forces in its immediate environment may have influenced

both the slow divergence of organisms and rapid mutations in contexts such as cancer.

Methods

smFRET based single-molecule DNA looping assay

Templates were purchased (IDT DNA) and converted into loopable molecules with 10 bp complementary

overhangs on either side, Cy3 and Cy5 fluorophores at the ends, and a biotin molecule (Supplementary

Note 1) via PCR amplification with KAPA Hi Fi Polymerase (Roche) and nicking near the ends by the site-

specific nicking enzyme Nt.BspQ1 (NEB). Molecules were immobilized on a PEG-coated quartz surface

(JHU slide production core for microscopy) functionalized with a small amount of biotin-PEG, via a

streptavidin sandwich, as described previously6. Immobilized molecules were incubated with T2.5 (2.5 mM


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NaCl, 10 mM Tris-HCl pH 8) for 1.5 hours. Low salt imaging buffer (20 mM Tris-HCl pH 8, 3 mM Trolx,

0.8% dextrose, 0.1 mg/ml glucose oxidase, 0.02 mg/ml catalase) was flowed into the channel and the

molecules were imaged on a TIRF microscope to determine the initial histogram of FRET values. High salt

imaging buffer (1 M NaCl, and all components of the low salt imaging buffer) was then introduced into the

channel at time 0, and FRET histograms were measured at various time points as done previously6. The

plot of the percentage of molecules with both donor-acceptor pairs in high FRET as a function of time was

fit to an exponential. Its time constant was defined as the looping time. The inverse of this was defined to

be the looping rate.

Loop-seq

Instead of individual templates, entire libraries representing as many as ~90,000 individual DNA sequences,

with the central 50 bp variable and flanked by identifcal 25 bp adapters, were obtained (Genscript), and

amplified using KAPA Hi Fi polymerase (Roche) in 20 cycles of emulsion PCR35 (ePCR) using the

Micellula DNA emulsion and purification kit (CHIMERx). The manufacturer’s guidelines were followed

during ePCR. ePCR prevents improper annealing among different template molecules via the common

adapter sequences. Amplification converted the library into 120 bp duplex molecules with a biotin near one

end, and the recognition sequence for the nicking enzyme Nt.BspQ1 (NEB) near both ends (Supplementary

Note 1). 20 µl of streptavidin-coated magnetic beads (Dynabeads MyOne Streptavidin T1, Thermo Fisher

Scientific) were washed 2x with 400 µl T50 BSA (1 mg/ml BSA (Invitrogen) in T50 (50 mM NaCl, 10 mM

Tris-HCl pH 8.0)) and resuspended in 20 µl T50 BSA. 2 µl of ~4 ng/ul amplified DNA was mixed with 5

µl of water, and 20 µl of the washed magnetic beads were added. After incubation for 10 minutes, the DNA

bound beads were washed 2x with 200 µl T50BSA and 1x with 200 µl T10BSA (1 mg/ml BSA (Invitrogen)

in T10 (10 mM Tris-HCl pH 8.0, 10 mM NaCl)). Digestion mix (84 µl water, 10 µl 10x NEB Buffer 3.1, 6

µl Nt.BspQ1 (NEB)) was prepared and heated to 50 oC for 5 minutes. Digestion resulted in an immobilized

library, where every DNA molecule has a central 50 bp duplex variable region, flanked by 25 bp left and

right adapters and 10 nt complementary single-stranded overhangs (Fig. 1d, Supplementary Note 1). The


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beads were pulled down and incubated with the heated digestion mix for 25 mins at 37 oC. The beads were

then washed 2x with 100 µl of T10BSA preheated to 50 oC, followed by 200 µl of T2.5BSA (1 mg/ml BSA

(Invitrogen) in T2.5 (10 mM Tris-HCl pH 8.0, 2.5 mM NaCl)). The beads were incubated in 200 µl

T2.5BSA for 1.5 hours on a rotor at room temperature. The bead sample was then split into two 95 µl

fractions denoted ‘sample’ and ‘control’. The beads in the sample fraction were pulled down and

resuspended in 200 µl looping buffer (1M NaCl, 1 mg/ml BSA, 10 mM Tris-HCl pH 8) for 40 seconds.

High salt (1M NaCl) initiates looping, which allows the complementary single-stranded overhangs at the

ends to stably hybridize6. Apparent DNA bendability has been shown to be independent of the salt

concentration used6.The tube containing the sample was then placed on magnets for an additional 35

seconds. The looping buffer was replaced with 200 µl of digestion buffer (6.66 µl of RecBCD (NEB), 20

µl NEB 10X Buffer 4, 20 µl of 10x ATP (NEB), 154 ul water) for 20 minutes. This was defined as looping

for 1 minute. In general, looping for n minutes implies incubation in looping buffer for up to 20 seconds

prior to the completion of n minutes, followed by 35 seconds over magnets before the solution was replaced

with digestion buffer. After 20 minutes, digestion buffer was removed by pulling down the beads and

replaced with 200 µl of looping buffer. The control was treated in exactly the same way, except the digestion

buffer had 6.66 µl of water instead of RecBCD. Beads in the sample and control fractions were then pulled

down and the looping buffer was replaced with 50 µl of PCR mix (25 µl 2x HiFi KAPA Hot Start ready

mix (Roche), 1 ul each of 100 µM primers (supplementary note 1), 23 µl water) and PCR amplified (16

cycles). If the library contained less than 20,000 sequences, the products were sequenced on an Illumina

MiSeq machine. For more complex libraries a HiSeq machine was used. Library preparation for sequencing

was done using the Nextera XT primer kit and followed a protocol similar to the Illumina protocol for 16S

metagenomic sequencing library preparation.

Sequencing results were mapped to the known sequences in the library using bowtie36. The number

of times each sequence was represented in the sample and control was obtained and 1 was added to all

counts. The relative population of each sequence in the digested and control pools was calculated.

Cyclizability of a sequence was defined as the natural logarithm of the ratio of the relative population of a


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sequence in the sample pool to that in the control. In addition to Bowtie36, Samtools37 and MATLAB

(Matworks) were used to analyze the data.

Purification of INO80:

INO80 was purified according to a protocol published earlier38. Briefly, S. cerevisiae cells were grown in

12 liters of YPD medium to an O.D. of 1.5. Frozen yeast cells were lysed in a SPEX freezer mill (15 cycles:

precool 2 min, run time 1 min, cool time 1 min, rate 15 cps). Ino80-3Flag was affinity-purified from whole

lysate using anti-Flag M2 agarose beads and eluted with Flag peptide (0.5 mg/ml). The complex was further

purified by sedimentation over a 20-50% glycerol gradient. Peak INO80 fractions were pooled and

concentrated using Centricon filters (50 kDa cut off), and buffer changed to 25 mM HEPES–KOH (pH

7.6), 1 mM EDTA, 2 mM MgCl2, 10% glycerol, 0.01% NP-40, 0.1 M KCl. Aliquots of purified INO80

were flash-frozen and stored at −80°C. Recombinant INO80 was also purified as per earlier protocols21.

Nucleosome sliding by INO80:

Nucleosome preparation and sliding by INO80 was performed under conditions as reported earlier19.

Sliding for in the presence of various [INO80] as reported in Fig. 2f (for 1 minute) and Extended Data Fig.

7 (for 1 minute and 2.5 minutes) was performed in 10 µl reaction volumes containing 8 nM nucleosomes

(nucleosomes formed on both constructs in the pair were present in equimolar proportion), 2 mM ATP, 24

mM tris-HCl pH 7.5, 43 mM KCl, 2.86 mM MgCl2, 0.55% glycerol and indicated concentration of INO80.

The mixture was incubated without ATP at 30 oC for 7 minutes. After addition of ATP, the reaction was

allowed to proceed for 1 minute at 30 oC, and was then quenched by the addition of lambda DNA and ADP

to final concentrations of 66.7 µg/ml and 20 mM respectively. For all sliding experiments reported in

Extended Data Fig. 6b (timecourse of INO80 sliding), conditions were the same except incubations prior

to ATP addition and the subsequent sliding reaction were carried out at room temperature. The reaction

was continued for the indicated amounts of time in presence of saturating INO80 prior to quenching.


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Quenched reactions were loaded on to 6% TBE gels (Invitrogen) in presence of 10% glycerol and run at

150 V for 1.5 hrs. The gel was imaged separately for Cy3 and Cy5 fluorescence.

Figure 1: A high-throughout method to measure the mechanical properties of DNA. (a) Schematic of

the single-molecule looping assay, showing how looping results in an increase in FRET. (b) In situ nicking

of 120 bp duplex DNA 10 nt from either end, following by washing with buffer at 50 oC, results in 100 bp

duplex molecules flanked by 10 nt single-stranded overhangs. Site-specific nicking is achieved by

engineered recognition sequnces of the nicking enzyme Nt.BspQ1 (NEB). See Supplementary Note 1. (c)

Percentage of DNA molecules in the high FRET (looped) state as a function of time after adding high salt,


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as observed for 10 different DNA sequences (Supplementary Note 1). Only molecules that contained both

the FRET donor and acceptor were considered. Solid lines represent exponential decay fits, whose

characteristic time constant is defined as the looping time and is listed for the 10 sequences in the bar plot

(Supplementary Note 1). (d) Schematic of a typical DNA fragment in a library just prior to looping. ‘n’ is

the distance in nucleotides of the biotin tether from the end of the molecule (e) Schematic of loop-seq

performed on a hypothetical library comprising only two sequences – green and pink. The library is

amplified via emulsion PCR35, which prevents improper annealing among different library members via

the common adapter sequences at the ends. The amplified library is immobilized on beads via biotin-

streptavidin interactions and nicked in situ to generate loopable molecules (Fig. 1d). After looping for 1

minute in high salt followed by the digestion of unlooped molecules and the amplification of surviving

molecules, the relative populations of green and pink in the digested fraction are 2/3 and 1/3 respectively.

The corresponding values in the control fraction are 1/2 and 1/2. The cyclizability of green is thus

𝑙𝑜𝑔! $"/$%/"% and that of pink is 𝑙𝑜𝑔! $

%/$%/"% . (f) Left panel: Cyclizabilities of 10 sequences (listed in

Supplementary Note 1) which were part of the Cerevisiae Nucleosomal Library (Supplementary Note 4) as

obtained by loop-seq vs looping times of those 10 sequences obtained from smFRET experiments (Fig. 1c).

Right panel: Violin plot of the cyclizabilities of all 19,907 sequences in the Cerevisiae Nucleosomal

Library. Cyclizabilities of the 10 sequences whose looping times were measured via smFRET are indicated

by ‘x’.


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Figure 2: DNA mechanics contributes to nucleosome depletion at the NDR and modulates remodeler

activities. (a) Schematic of the Tiling Library. The central 50 bp variable region was designed to tile 576

genes in S. cerevisiae at 7 bp resolution from 600 bp upstream till 400 bp downstream of the dyads of the

+1 nucleosomes of these genes every 7 bp (Supplementary Note 9). DNA is represented by the blue lines,

and nucleosomes by the oval shapes. (b) Mean intrinsic cyclizability (with (black) and without (grey) any

smoothening) and nucleosome occupancy vs position from the canonical location of the dyad of the +1

nucleosome, averaged over all 576 genes in the Tiling Library. Nucleosome occupancy values17 and

positions32 were as reported earlier. All genes are oriented in the direction of transcription. The blue dashed


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line (at -73 bp) marks the upstream edge of the +1 nucleosome. The red dashed line marks the start of the

rigid DNA region, approximated as the midpoint between the two red arrows. See supplementary note 10.

(c) Intrinsic cyclizability and nucleosome occupancy vs position from the dyads of the +1 nucleosomes of

two individual genes (see Extended Data Fig. 5 for six more examples). (d) Schematic showing an INO80

enzyme attempting to slide a +1 nucleosome upstream of its canonical location. The rigid DNA region starts

~40 bp upstream of the +1 nucleosome edge (Fig. 2b), about where the Arp8 module of an INO80

attempting to slide the nucleosome upstream would bind21,23. (e) Schematic of the experiment comparing

the effect of DNA rigidity on nucleosome sliding by INO80. A pair of constructs were engineered involving

a nucleosome formed on the 601 DNA sequence attached to two different 80 bp linker DNAs that had a

large difference in intrinsic cyclizability values around the middle of the 80 bp region. The construct with

the lower intrinsic cyclizability (i.e. more rigid) 80 bp region was labeled with Cy3 at the 5’ terminus distal

to the 601 region and the other construct was labeled with Cy5. Both constructs were present in the same

reaction volume, and sliding was effected for 1 minute in the presence of INO80 and ATP. The products

were run on a gel. Sliding results in the generation of centered nucleosomes, which migrate more slowly23.

The gel was imaged separately for Cy3 and Cy5 fluorescence to quantify the extent of sliding of

nucleosomes formed on the two constructs in the pair independently. See Supplementary Note 11. (f) Three

separate nucleosome sliding experiments were performed involving three different pairs of nucleosome

constructs as described in figure 2e and supplementary note 11. In each experiment, four concentrations of

INO80 were used (in lanes L1 – L4): 2, 6, 9, and 13 nM. The products of the sliding reaction were run along

a 6% TBE gel, which was imaged separately for Cy3 and Cy5 fluorescence. For each concentration of

INO80, i.e. for each lane, the nucleosome sliding extents in the two constructs in the pair were quantified

(bar plots) as described in figure 2e and supplementary note 11.


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Figure 3. DNA mechanics contributes to global nucleosome organization. (a) Mean intrinsic

cyclizability and nucleosome occupancy vs position from the dyad of the +1 nucleosome, averaged

separately over 185 and 345 genes in the tiling library that have been classified as possessing stable and

fragile -1 nucleosomes respectively24. Plots were generated as in figure 2b. (b) Mean intrinsic cyclizability

and nucleosome occupancy vs position from the +1 nucleosomal dyad, averaged over 227 identified genes

along S. cerevisiae chromosome V. Grey background represents s.e.m. See supplementary note 13. (c) Top:

intrinsic cyclizability vs position from the dyad, averaged over all 3,192 nucleosomes in chromosome V.

Shaded background represents s.e.m. Bottom: Nucleosomes were sorted into quartiles based on reported

NCP scores32, and average flexibility profiles across nucleosomes were plotted for each quartile. See


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18

Extended Data Fig. 8a-b for quartile boundaries, and for plots along octiles. (d) Intrinsic cyclizability and

nucleosome occupancy vs position along all genes in chromosome V (blue) and among the 34% of these

genes that had the highest (red) and lowest (green) +1 nucleosome NCP scores32. Plots were obtained as in

panel b. The three vertical dashed lines mark the edges and dyad of the +1 nucleosome. Shaded backgrounds

represent s.e.m. (e) Mean (solid line) and s.e.m. (grey background) of intrinsic cyclizability values around

the dyads of gene body nucleosomes that lie between the TSS and the TTS of all identified 227 genes along

chromosome V in S. cerevisiae. (f) Mean and s.e.m. of intrinsic cyclizability values of DNA in a 50 bp

window around the dyads of gene body nucleosomes (blue) and in a 50 bp window around the edges of

gene body nucleosomes (from positions -73 till -56 and from +56 till +73 with respect to the dyad) (red).

Solid lines are guides to the eye. (g) Left of dashed line: intrinsic cyclizability as a function of position

along the native sequences of the 500 +7 nucleosomes represented in Library L (supplementary note 14).

Right of dashed line: same quantity for the four sets of codon-altered sequences. The first two plots are for

the cases where synonymous codons were selected considering the natural codon-usage frequency in S.

Cerevisiae. This was not the case for the subsequent two plots. Solid line represents mean, and the grey

background represents s.e.m. Data were plotted with a rolling window averaging of 7 adjacent fragments.


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Figure 4: TSS-proximal nucleosomes are asymmetric. (a) Black: Intrinsic cyclizability as a function of

position along 601 DNA (supplementary note 18) Blue: abscissa value of the solid horizontal line (-0.196)

is the mean intrinsic cyclizability along the 500 native +7 nucleosomal sequences represented in Library L

(supplementary note 14). The height of the light blue background (0.011) is twice the s.e.m. (b) Schematic

representing the design of the Cerevisiae Nucleosomal Library (supplementary note 4). The library contains

DNA fragments taken from the 50 bp immediately to the left and right of the dyads of the ~10,000

nucleosomes in S. cerevisiae that have the highest NCP scores. (c) Mean intrinsic cyclizabilities of the 50

bp DNA fragments that lie immediately adjacent to the TSS proximal (red) or distal (blue) side of the dyads

of various categories of nucleosomes (-9 through +9) (see supplementary note 17). Error bars are s.e.m. (d)

A subset of the data as in panel c, where the means were calculated considering only genes among the 10%

most (left panel) or least (middle panel) expressed in S. cerevisiae. The right panel is identical to panel b,

except for an altered y-axis scale. Error bars are s.e.m. See supplementary note 17.


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Author Contributions: AB and TH designed research. AB performed research and analyzed data. AB and

TH wrote the paper. Other authors contributed in the following areas: DGB – extraction of nucleosome

occupancy from published data and plectoneme density calculations. ZQ – preparation of some libraries.

TK, TN – initial assay development and the characterization of RecBCD. AR, SE, KPH – purification of

INO80 and related insights. BC – observation of the phasing effect. MTM, CW – initial development of

nucleosome assays. MH, TR, JSS – initial analysis of sequencing data. All authors commented on the

manuscript.

Competing Interests: The authors declare no competing interests.

Data Availability: All raw data and accession codes will be made available upon request.

Code availability: All codes used to analyze sequencing data will be made available upon reasonable

request.

Acknowledgements: A.B. and T.H. would like to thank Carl Wu, Xinyu Feng, and Matthew F. Poyton for

insights and help related to INO80 biochemistry, and Quicen Zhang for help with initial assay development

efforts. K.P.H and S.E. would like to thank Manuela Moldt for purification of recombinant INO80. This

work was supported by the National Science Foundation Grants PHY-1430124 and EFMA 1933303 (to

T.H.), the National Institutes of Health Grants GM122569 (to T.H.), the National Institutes of Health grant

NIH R01CA163336 (to J.S.S.), the European Research Council (Advanced Grant INO3D to K.P.H), and

by the Deutsche Forschungsgemeinschaft (CRC1064 and Gottfried Wilhelm Leibniz-Prize to K.P.H). A.B.

was a Simons Foundation Fellow of the Life Sciences Research Foundation. T.H. is an Investigator with

the Howard Hughes Medical Institute.

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